Electrophysiological measurements of synaptic connectivity and plasticity in the longitudinal dentate gyrus network from mouse hippocampal slices

Summary Longitudinal synaptic connections between dentate gyrus (DG) granule neurons in the hippocampus have been found to be correlated with increased anxiety. Here, we present a protocol to assess synaptic connectivity and plasticity in the longitudinal DG network. We detail the steps for (1) obtaining acute mouse hippocampal slices that contain longitudinal DG-DG connections, (2) measuring excitatory postsynaptic potentials using whole-cell patch clamp recording combined with two-photon microscopy and glutamate uncaging, and (3) assessing synaptic plasticity using extracellular field recording. For complete details on the use and execution of this protocol, please refer to Pak et al. (2022).1


Highlights
Obtain healthy hippocampal slices that contain intact DG-DG longitudinal synapses Measure postsynaptic membrane potential in whole-cell patched DG granule cells (DGGCs) Induce EPSPs in patched DGGCs using two-photon microscopy and glutamate uncaging Measure synaptic plasticity of DG-DG synapses using extracellular field recording SUMMARY Longitudinal synaptic connections between dentate gyrus (DG) granule neurons in the hippocampus have been found to be correlated with increased anxiety. Here, we present a protocol to assess synaptic connectivity and plasticity in the longitudinal DG network. We detail the steps for (1) obtaining acute mouse hippocampal slices that contain longitudinal DG-DG connections, (2) measuring excitatory postsynaptic potentials using whole-cell patch clamp recording combined with two-photon microscopy and glutamate uncaging, and (3) assessing synaptic plasticity using extracellular field recording. For complete details on the use and execution of this protocol, please refer to . 1

BEFORE YOU BEGIN
All animal handling procedures were approved by the Institutional Animal Care and Use Committee of City University of Hong Kong (A-0117) and Incheon National University (INU-ANIM-2017-08).
Before the day of the experiment Timing: 2 h CRITICAL: Avoid soap, detergent, ethanol, and other chemicals when washing equipment, as they may interfere with physiology.

Prepare a holding chamber.
Optional: A slice holding chamber can be crafted using a small food container (85 mm diameter), nylon stockings, plastic mesh sheet, 60 mm diameter petri dish, silicone tubing, tubing connectors, and a hot glue gun ( Figure 1B). The nylon stockings, plastic mesh sheet and petri dish are assembled into an inner chamber. The inner chamber is inserted into the small food container and should be secured by hot gluing it to the bottom of the food container. It should also be surrounded by the plastic mesh sheet barrier to prevent big air bubbles from getting inside the inner chamber. The silicone tubing has small holes made with a 27G needle to allow carbogen (95% O 2 /5% CO 2 ) to bubble into the outer chamber, and is placed along the outer periphery of the inner chamber. During electrophysiological recording, this inner chamber is kept submerged in ACSF or slicing medium and the brain slices are placed on top of the nylon ( Figure 2J).

Configure stimulation protocols on [Clampex] and [Prairie View
] software for whole-cell patch recording and extracellular field recording according to the manufacturers' instruction manuals. a. Protocol to deliver 100 ms of increasing current to a whole-cell patched neuron every 10 s for 5 min (step 36). b. Protocol to determine the optimal number of stimulation spots, laser duration, stimulation power and dot size for glutamate uncaging (step 47). Recommended to apply minimum power for 1 ms and increase power incrementally every 30 s. c. Protocol to determine the half-maximal stimulation amplitude for extracellular field recording (steps 70-74) by increasing stimulation amplitude at 30 s intervals. d. Protocol to deliver high-frequency stimulation (100 Hz, 1 s, 4 times at 1 s intervals) (step 77).
On the day of the experiment Timing: 45 min 4. Prepare solutions and bench: a. Pull a glass capillary to make micropipettes using a puller.
Note: A glass capillary with an inner diameter of 1.65 mm is recommended. Target the tip resistance at 5-7 MU for DGGC whole-cell patch recording and 3-4 MU for extracellular field recording.
b. Pre-chill the experiment tools (such as the dissection tools and the vibratome parts, i.e., specimen disc, buffer tray, ice tray) at À20 C for at least 15 min ( Figure 1). CRITICAL: The atmospheric temperature around the dissection area should be between 18 C-20 C.
c. Add MgCl 2 .6H 2 O and CaCl 2 .2H 2 O from its stocks to 400 mL ACSF and 400 mL slicing medium. The volume may vary depending on the length and setting of your experiment.
d. Pre-oxygenate the slicing medium on ice with carbogen using an air stone for at least 20 min. e. For whole-cell patch recording, pour approx. 200 mL slicing medium into the holding chamber, and place the entire holding chamber inside a water bath set to 32 C.  f. For extracellular field recording, pour approx. 200 mL ACSF into the holding chamber and keep it at room temperature (18 C-20 C).
Note: Adjust the amount of solution in the holding chamber according to the size of the chamber. There should be enough solution inside to submerge the slices.
CRITICAL: Oxygenate the ACSF holding chamber for at least 20 min before starting dissection. Adjust the gas pressure so that the ACSF fizzes lightly, ensuring that no big bubbles form under the nylon of the inner chamber.
5. Prepare dissection and slicing tools ( Figure 1A): a. For example plastic bag, dissection tools (decapacitation scissors, spring scissors, forceps, scalpel), two stainless steel spatulas (one with a curved end), stainless steel spoon (1.5 cm diameter), fine brush, super glue. b. Cut the tip of a disposable 5 mL pipette diagonally to use it for transferring brain slices. c. On a flat cold pack (around 17 3 25 cm), place a petri dish with filter paper inside. The filter paper helps to prevent the tissue from slipping during dissection. d. Stick a piece of masking tape on the specimen disc. The brain tissue will be superglued onto the tape. 6. Chill the slicing medium for 10-15 min at À80 C until it is semi-frozen. 7. Assemble the vibratome ( Figure 1B): a. Secure the buffer tray inside the ice tray. b. Fill the ice tray with ice and water while preventing any from getting inside the buffer tray. c. Attach the ice tray to the vibratome. d. Attach the blade holder and blade as in Figure 1C. Set the clearing angle at 18 (effective clearance angle is 3 ). The blade is added at the last moment before beginning the slicing procedure. e. Lower the blade into the buffer tray. 8. Oxygenate the semi-frozen slicing medium and scrape the frozen crystals from the bottle walls with a spatula. Shake the bottle vigorously to make a slurry. 9. Fill the buffer tray with approx. 150 mL slicing medium slurry. Pour approx. 40 mL into the beaker (inside the ice box), and the remaining 10 mL into the petri dish (on top of the cold pack). 10. Oxygenate the slicing medium in the buffer tray.
Note: You may need to wait until there are fewer ice crystals inside the buffer tray before proceeding with the dissection (Figure 1C), as having too many crystals may damage the brain tissue. Recently, dentate gyrus granule cells (DGGCs) were discovered to target adjacent DGGCs 3-5 via longitudinal axons that span across the dorsoventral (or septotemporal) axis of the hippocampus. [6][7][8] Unlike transverse axons along the DG-CA3 network, these DG-DG axons are involved in anxiety-like behavior. 1

KEY RESOURCES
Steps 1-7 detail the steps for obtaining acute longitudinal dentate gyrus slices in order to measure the extracellular excitatory postsynaptic potentials (fEPSPs) and examine the synaptic transmission and plasticity of longitudinal DG-DG synapses.
1. Anaesthetize a 3-4-week-old mouse by isoflurane inhalation in a closed chamber inside a fume hood. 2. When its breathing slows to around one beat per sec, decapacitate with scissors at the cervicothoracic junction to remove the spinal cord from the base of the skull. 3. Isolate the whole brain with dissection tools, being careful not to touch the hippocampus.
a. Cut the skin from the base of the skull toward the nose using standard scissors. Peel and cut away the skin on the top and sides to completely expose the skull. b. Make lateral cuts on both sides of the vertebrae and cut through the jaw bones. c. Insert one blade of the microscissors under the skull and cut towards the eyes along the midline. At the bregma line, make two lateral cuts on the skull from the midline towards the eyes. d. Use forceps to open the skull from the incision and expose the whole brain.
CRITICAL: Avoid applying pressure onto the brain while removing the skull.
e. Insert the thin, flat end of the spatula near the olfactory bulb and scoop up the brain from the ventral side. Gently cut the optic and cranial nerves with the spatula to free the brain from the skull. f. Gently submerge the brain into the 50 mL beaker containing the slicing medium for 20 s on ice.
This hardens the tissue, which facilitates slicing. 4. Dissect out the hippocampus and transfer it to the vibratome.
a. Transfer the brain onto the cold petri dish. Use a stainless-steel spoon to transfer the brain, ensuring that only the rostroventral side of the brain comes in contact with the spoon. b. Cut out the cerebellum using a scalpel. c. Separate the left and right hemispheres with a clean top-down cut using a scalpel ( Figure 2C). d. Insert the blunt end of one spatula into the gap between the cerebral cortex/hippocampus and the midbrain ( Figure 2D). Use this to hold down the tissue but avoid pressing the spatula onto the cerebral cortex/hippocampus. e. Use the other spatula to cut away the midbrain, thalamus, and septum. This will expose the 'pocket' where the hippocampus is located. f. Gently isolate the hippocampus from the cerebral cortex using small, scraping motions with the blunt end ( Figure 2E). g. Cut out the septal (medial) and temporal ends (lateral) of the hippocampus using a scalpel ( Figure 2F).
Note: Cut the septal and temporal ends at different angles to help you recognize the orientation at later steps.

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h. Spread a thin, even layer of superglue on the masking tape. Use just enough glue to cover the size of the block of the hippocampus (approx. 3 3 3 mm). i. Transfer the hippocampus onto the curved side of the spatula using a fine brush. Touch the brush on a piece of tissue paper to wick away any excess solution from the brain. j. To produce longitudinal slices, stick the CA3 (rostral) side on the glue (Figures 2G and 2H).
This will orient the tissue so that the blade cuts parallel to the dorsoventral/septotemporal plane. k. Submerge the disc inside the buffer tray.
Note: Orient the disc so that the tissue is sliced from the septal to the temporal end (or vice versa) rather than from CA1 to DG (or vice versa) ( Figure 2I). 5. Produce hippocampal slices using the vibratome.
a. Set the thickness to 300 mm (whole-cell patch recording) or 400 mm (extracellular field recording). b. 0.05 mm/s speed and 1.20 mm amplitude are recommended. c. Cut away the top 500-600 mm of tissue (depending on the age of the animal), and slice until the CA1 and DG are visible. When visualized under light, the CA1 is clearly visible with bare eyes.
Note: The best longitudinal slices will contain one layer of CA1 and two layers of DG when observed under a light microscope ( Figure 2J). One block of the hippocampus may produce 2-3 slices.
6. Using a blunted disposable pipette, gently but quickly transfer the slices into the inner chamber of the oxygenated holding chamber containing ACSF or slicing medium ( Figure 2J).
CRITICAL: Always keep the slices submerged. Prevent the inner chamber from floating up. Remove any air bubbles from the inner chamber using a pipette, but avoid touching the slices.
Note: The middle part of the chamber provides the best oxygenation for slices.

Recover slices
Timing: 60 min 7. Let the brain slices rest inside the holding chamber, which is to be continuously oxygenated. a. For whole-cell patch recording, place the entire holding chamber inside a 32 C water bath for 30 min, then transfer it to room temperature to rest for an additional 30 min minimum. b. For extracellular field recording, rest the brain slices inside the holding chamber at room temperature for at least 1 h.

Whole-cell patch clamp
Timing: 1 h Steps 9-32 help achieve whole-cell patch recording of DG granule neurons (DGGCs) in longitudinal DG slices. Subsequently, focal glutamate uncaging on the dendrites of neighboring DGGCs will elicit excitatory postsynaptic potentials (EPSPs) in the patched cells (steps 33-55). EPSPs will be recorded from patched DGGCs.
Note: DGGCs in slices may survive for up to 5 h after dissection.
9. Thaw an aliquot of internal solution on ice. 10. Configure the following: a. On one arm of the micromanipulator, mount the Current Clamp and Voltage Clamp Headstage. Then attach the silver chloride recording electrode/pipette holder (do not fit the glass micropipette yet). b. Adjust the angle of the Current Clamp and Voltage Clamp Headstage to 15 from the horizontal plane. c. Attach 1 mm diameter tubing (approx. 50 cm long) to the suction port of the pipette holder.
At the other end of the tubing, attach a 1 mL syringe connected via a three-way valve. d. Insert a silver chloride reference electrode into the bath solution without touching the brain slice. Then ground the reference electrode. 11. Add picrotoxin to the ACSF perfusion solution (final concentration at 50 mM). 12. Perfuse the specimen stage with oxygenated ACSF maintained at 31 C-32 C by an automatic temperature controller, supplied and discharged at a steady rate (1.5 mL/min) using peristaltic pumps.
CRITICAL: Check frequently to make sure that the stage does not flood and that there are no excessive vibrations or fluid currents.
13. Gently transfer one brain slice to the specimen stage with a blunted disposable pipette. Place a harp on top of the slice to secure its position.
Note: For glutamate uncaging, the laser path should not intersect with the pipette. Mind the orientation of the slice on the specimen stage so that the laser path can reach the molecular layer of the DG blade, but it does not contact the pipette and its path during whole-cell patch recording.
14. Acclimatize the slice on the specimen stage for an additional 30 min. 15. Visualize one slice with a 103 air objective under bright-field illumination. Check that the fluid level is stable and that there are no vibrations that could interfere with the patch process. 16. Switch to a 403 water immersion objective. Locate the dorsal blade (the DG layer closer to CA1).

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Note: DG granule cells can be identified by the densely packed arrangement of their soma into rows, which appear darker in color under bright-field illumination ( Figure 4B).
17. Switch to a water immersion 403 objective and find a healthy cell to patch. Mark the position of the cell with masking tape on the computer monitor. CRITICAL: Keep the pipette tip clean by not touching it with anything but the brain slice.  Figure 3C). a. Check that the tip is not broken and that there are no debris or air bubbles inside. 27. Lower the pipette towards the slice while maintaining the tip in focus ( Figure 3D). Use a finer control mode on the micromanipulator and the fine adjustment knob of the microscope. 28. Once the target neuron and pipette tip are visible at almost the same Z-axis position, focus on the target neuron and slowly approach the pipette towards the cell. a. When they come into contact, the positive pressure inside the pipette should produce a slight dent on the cell surface. The current pulse shown on the [Membrane Test] should also start to decrease. 29. Cell attachment -Immediately apply gentle suction by pulling out the syringe plunger or via mouth pipetting. The cell is successfully attached when the tip resistance reaches at least 1-5 GU.
Note: Holding the membrane potential at -81 mV in [MultiClamp], which is the typical resting membrane potential of DGGCs, helps to counteract membrane hyperexcitability that may occur immediately after breaking into a patched cell. The seal should form within around 1 min.

Configuration of two-photon laser microscope
Timing: 15 min 33. Configure a two-photon laser microscope (e.g., Bruker Ultima In Vitro two-photon microscope) equipped with two lasers (e.g., Coherent Chameleon Ultra Ti: Sapphire), each of which is tuned to an excitation wavelength of 810 nm (for image acquisition) or 720 nm (for glutamate poststimulation) and modulated by an electro-optic modulator (or Pockel cell, Conotopics, M350).
Note: It may take approx. 10 min for the laser to warm up after turning on the power.

Two-photon imaging of a patched neuron and its neurites
Timing: 45 min Steps 34-43 allow high-resolution visualization of a single DG neuron that was patched in steps 9-32.
34. External light sources should be minimized in a darkened room, including any halogen light sources. 35. Switch to a 603 objective lens. 36. On [Clampex], run a pre-configured protocol to inject the threshold current (pA) once every 10 s for 5 min.

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a. The threshold current is determined as the lowest current level required to induce a single action potential. b. Constant threshold current (pA) is applied for 100 ms.
Note: This process helps to monitor neuronal viability and promotes the fluorescence dye to spread to distal dendrites for clear visualization. CRITICAL: Increase the power one by one to prevent photodamaging the brain slice with the high laser intensity.  Note: [Step Size (mm)] of 1-2 mm is recommended for good-quality stacked images.
42. Place the cell body in the middle of the [Image Window] and record its position P 1 = ðx 1 ; y 1 ; z 1 Þ under [Stage Control], so that the patched neuron may be located again later. To maintain a stable whole-cell configuration, the pipette position may need to be readjusted at least every 5 min to stay close to the cell body ( Figure 4B). 43. Z-stack images are automatically saved in a folder defined by the user. Alternatively, the brightness or contrast can be adjusted for individual images and later use [ImageJ] software to create a stack with specified images.

Timing: 2 h
Steps 44-52 describe the procedure for evoking an EPSP in the patched DG neuron by photostimulation of another DG neuron(s) located in the same DG layer (ventral or dorsal). Steps 53 and 54 repeat the procedure with cadmium chloride (CdCl 2 ) to confirm that the EPSP was evoked by the vesicular release of neurotransmitters from the photostimulated DG neuron.
44. Freshly prepare 500 mM MNI-caged-L-glutamate in ACSF solution and perfuse it into the recording chamber. Wait at least 3 min for the solution to diffuse entirely throughout the recording chamber (for a perfusion speed of 1.5 mL/min).
CRITICAL: Maintain a stable whole-cell configuration during the whole experiment.
45. Visualize the unstained brain slice without a halogen light source using Dodt Gradient Contrast (DGC), which improves the contrast of images. Note: A hippocampal longitudinal slice contains two distinct layers of dentate gyrus granule cells (GC), which are the dorsal and ventral blades of granule cell layers. 3 Therefore, target the molecular layer that is continued from the granule cell layer of the patched granule neuron. For example, do not apply photostimulation on the molecular layer that extends from the ventral blade of GC layer, if you have patched a neuron in the dorsal blade of GC layer ( Figure 4B).

Open [Mark Points] from [Image Window
] and configure the settings as follows: a. Be sure to optimize the laser duration, power, number of stimulation spots, and dot size beforehand to ensure that they can elicit an action potential in a neuron. i. Start by applying the minimum power (10 mW) for 1 ms laser duration and increase incrementally.
ii. Rest the slices for 30 s between stimulations to avoid hyperexcitation and to give time for presynaptic neurons to recover neurotransmitter vesicles. b. 3 3 3 grid of point stimulation (5 ms laser duration, 10 mm spot spacing, and various interpoint delays, > 0.12 ms) at an average power of 42 mW (ranging from 7.5 to 78.08 mW) was used previously 3 ; the spot size ranged from a diffraction-limited spot of 0.  Figure 4B). The approximate distance between two points P 1 = ðx 1 ; y 1 ; z 1 Þ and P 2 = ðx 2 ; y 2 ; z 2 Þ in a three-dimensional space is calculated by the following generalized distance formula: To confirm that the EPSP was evoked from presynaptic vesicular release, add 0.1 mM cadmium chloride (CdCl 2 ) to the perfusate and wait a few minutes for it to diffuse entirely throughout the recording chamber. 54. Repeat steps 45-50 at the position P 2 and record EPSPs under the effect of CdCl 2 .
Note: CdCl 2 blocks chemical synaptic transmissions at presynaptic terminals. If the photostimulated neurons (presynaptic neurons excited by uncaged glutamate) have synaptic connections to the patched neuron (postsynaptic neuron), the peak amplitude of EPSPs of the patched neurons should decrease or disappear after the application of CdCl 2 . As the effects of CdCl 2 are reversible, recovery of EPSPs can be observed if steps 45-50 are again repeated without CdCl 2 . 55. After the experiment, analyze the peak amplitudes of all EPSPs offline using [Clampfit] software.

Extracellular field recording
Timing: 45 min Extracellular field recording using longitudinal DG slices allows assessment of the long-term synaptic plasticity of the DG-DG network. Steps 56-69 help to prepare brain slices for extracellular field recording.
56. After producing 400 mm-thick brain slices, set up the electrophysiology equipment as follows: a. On one arm of the micromanipulator, secure the concentric bipolar microelectrode (stimulating electrode) that is connected to the electrical stimulus isolator. b. On the other arm, mount the Current Clamp and Voltage Clamp Headstage (recording electrode headstage, angled at 20 ). Backfill the pulled capillary pipette (recording electrode) with ACSF and mount the glass micropipette to the Headstage. 57. Add picrotoxin to the ACSF perfusion solution (final concentration at 50 mM). 58. Perfuse the specimen stage with oxygenated ACSF maintained at 31 C-32 C by an automatic temperature controller, supplied and discharged at a steady rate (1.5 mL/min) using peristaltic pumps. a. Noise range within 0.2 mV is acceptable. If too high, refer to troubleshooting -problem 2). 65. Using the micromanipulator, lower the stimulating and recording electrodes to hover just above the slice. 66. Lower the stimulating electrode to lightly come into contact with the inner molecular layer (IML) of the dorsal blade of DG granule cell layer ( Figure 5). 67. Lower the recording electrode to touch the same IML of the dorsal blade of DG granule cell layer 200-400 mm away from the stimulating electrode. 68. Evoke local field potentials (LFPs) with a minimum constant current at 100 ms duration and repeat at 30-s intervals. 69. In the meantime, adjust the position of the recording electrode to find the area that gives maximum amplitudes and an exemplary shape of LFPs (for reference, see Pak et al. 1 ). a. If no responses are detected, change the location of the recording electrode, or use another brain slice.

Timing: 20 min
Steps 70-74 help determine the half-maximal stimulation amplitude which is the stimulation intensity that will be applied for assessing synaptic plasticity in steps 75-79. 70. Configure the digitizer, the pulse stimulator, and the electrical stimulus isolator. See the Axon Guide by Molecular Devices for information on how to set up the various equipment. Running a protocol in [Clampex] triggers the [Master-9] pulse stimulator to control the stimulus isolator to stimulate through microelectrode.

EXPECTED OUTCOMES
For a whole-cell patched DG granule cell filled with Alexa Fluor 594, its cell morphology will be visible under a two-photon laser scanning microscope. Its soma, dendrites, and axons will be visible, particularly the longitudinal axons that project to neighboring DG neurons on a longitudinal DG slice. Then, EPSPs can be recorded from whole-cell patched DGGCs that are synaptically activated by a group of DGGCs excited by uncaged glutamate. On longitudinal DG slices, long-term synaptic potentiation can be induced and observed by measuring extracellular field EPSPs from the molecular layer of the DG. These expected results suggest the existence of longitudinal DG-DG connections that possess long-term synaptic plasticity.

LIMITATIONS
It is still uncertain whether the DG-DG synaptic response in longitudinal slices solely involves DG granule neurons. Mossy cells may contribute to the recorded responses.
These methods cannot identify whether the long-term potentiation of DG-DG connections involves presynaptic mechanisms, postsynaptic mechanisms, or both.

Potential solution 1
The most important factors for obtaining healthy slices are dissecting at cold temperatures, providing sufficient oxygen, dissecting as quickly as possible, and gentle handling of tissue.
Prepare new solutions. Check water quality, pH and osmolarity.
Optimize the recovery period. Shorten, lengthen, or even eliminate the period in the 32 C water bath or at room temperature. Optimize the vibratome settings. Check that the slices have an even surface and consistent thickness. Adjust the blade speed (e.g., 1.2 mm/s) and amplitude (e.g., 1.00 mm).

Potential solution 2
Unplug electronic devices connected to the rig one by one to find the noise source. Change the reference electrode. Ensure that the tip only touches the surface of the bath solution and no other surfaces of the electrophysiology set-up. Re-chloride the reference electrode. Adjust the capacitance and gain settings of the amplifier. Check the grounding of all components. For components that cannot be grounded, try shielding with aluminum foil connected to a ground box via wires and an alligator clip. The aluminum foil should not touch the table.
Clean the pipette holder and dry it thoroughly before use. Check that the bath solution is not leaking from the stage. Salt build-up around the bath imaging chamber, or the container collecting the solution from the perfusion outlet, can contribute to noise. Change the positions of electronic devices nearby the rig, e.g., wifi routers.

Problem 3
Failure to induce fEPSP in extracellular field recording (steps 72 and 73).

Potential solution 3
Check for the presence of fiber volley from fEPSP from the Scope window. If present, change the position of the electrodes, or adjust lowpass and highpass filters. If absent, check a different slice. Dead slices produce a fiber volley but no synaptic potentials. Prepare healthier brain slices.

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